Depletion of SMN protein in mesenchymal progenitors impairs the development of bone and neuromuscular junction in spinal muscular atrophy

  1. Sang-Hyeon Hann
  2. Seon-Yong Kim
  3. Ye Lynne Kim
  4. Young-Woo Jo
  5. Jong-Seol Kang
  6. Hyerim Park
  7. Se-Young Choi  Is a corresponding author
  8. Young-Yun Kong  Is a corresponding author
  1. School of Biological Sciences, Seoul National University, Republic of Korea
  2. Department of Physiology, Dental Research Institute, Seoul National University School of Dentistry, Republic of Korea

Abstract

Spinal muscular atrophy (SMA) is a neuromuscular disorder characterized by the deficiency of the survival motor neuron (SMN) protein, which leads to motor neuron dysfunction and muscle atrophy. In addition to the requirement for SMN in motor neurons, recent studies suggest that SMN deficiency in peripheral tissues plays a key role in the pathogenesis of SMA. Using limb mesenchymal progenitor cell (MPC)-specific SMN-depleted mouse models, we reveal that SMN reduction in limb MPCs causes defects in the development of bone and neuromuscular junction (NMJ). Specifically, these mice exhibited impaired growth plate homeostasis and reduced insulin-like growth factor (IGF) signaling from chondrocytes, rather than from the liver. Furthermore, the reduction of SMN in fibro-adipogenic progenitors (FAPs) resulted in abnormal NMJ maturation, altered release of neurotransmitters, and NMJ morphological defects. Transplantation of healthy FAPs rescued the morphological deterioration. Our findings highlight the significance of mesenchymal SMN in neuromusculoskeletal pathogenesis of SMA and provide insights into potential therapeutic strategies targeting mesenchymal cells for the treatment of SMA.

eLife assessment

This important work by Hann et al. advances our understanding of the role of the survival motor neuron (SMN) protein in coordinating pathogenesis of the spinal muscular atrophy (SMA). The authors addressed many concerns raised by the reviewers, providing convincing evidence in terms of skeletal analyses not being able to satisfactorily elucidate SMN regulation of bone development.

https://doi.org/10.7554/eLife.92731.3.sa0

Introduction

The survival motor neuron (SMN) protein is a crucial component of the spliceosome complex and is essential for the proper function of all cell types (Mercuri et al., 2022). Deficiency in SMN protein disrupts the formation of spliceosome complexes, ultimately causing splicing defects in multiple genes. Mutations in the SMN1 gene, which encodes the SMN protein, give rise to the neuromuscular disorder spinal muscular atrophy (SMA). SMA is characterized by neuromuscular junctions (NMJs) disruption, muscular atrophy, and alpha motor neuron loss (Mercuri et al., 2022; Burghes and Beattie, 2009). The severity of the disease in humans correlates with the copy number of SMN2, which is a paralog of SMN1 in humans. SMN2 primarily produces less functional exon 7-deleted SMN protein and rarely generates a limited quantity of functional full-length SMN protein via alternative splicing. SMA patients are classified into types 0 through 4 based on the severity and timing of disease onset, predominantly determined by the number of copies of the SMN2 gene they possess. More than 50% of SMA patients are categorized as type 1, characterized by muscle defects with proximal muscle atrophy during infancy, eventually resulting in death within a few years (Mercuri et al., 2022).

Previous studies suggest that the onset of SMA is mainly attributed to SMN loss in motor neurons (Monani et al., 2000; Burghes and Beattie, 2009). Nevertheless, motor neuron-specific SMN deficiency in the SMA mouse model exhibits relatively mild phenotypes compared to whole-body SMA mouse models (Park et al., 2010; McGovern et al., 2015). Furthermore, restoring SMN to motor neurons in SMA mouse models result in only partial rescue in lifespan and neuromuscular defects (Passini et al., 2010; Martinez et al., 2012; McGovern et al., 2015; Besse et al., 2020). Systemic administration of antisense oligonucleotide (ASO), which corrects SMN2 splicing to restore SMN expression, significantly prolongs survival compared to central nervous system (CNS) administration (Hua et al., 2011). In the mouse treated with a systemically delivered ASO, blocking the effect of ASO in the CNS by a complementary decoy did not have any detrimental effect on survival, motor function, or NMJ integrity (Hua et al., 2015). These studies suggest that peripheral SMN plays a crucial part in SMA pathology. Overall, investigating the impacts of SMN depletion in peripheral tissues is critical for alleviating neuromuscular impairments and increasing life expectancy in SMA.

In SMA patients, bone growth retardation has been observed (De Amicis et al., 2021; Kipoğlu et al., 2019; Hensel et al., 2020). Studies using whole-body SMA mouse models have revealed that this is caused by diminished growth plate chondrocyte density and endochondral ossification defects, independent of muscle atrophy (Hensel et al., 2020). However, it is still unclear whether these defects result from the SMN ablation in bone-forming cells, or from a decline in liver-derived insulin-like growth factor (IGF) in SMA patients and mice (Hua et al., 2011; Yesbek Kaymaz et al., 2016). In severe SMA mice, the serum levels of IGF decreased by approximately 60% or became undetected (Hua et al., 2011; Murdocca et al., 2012). The decreased serum IGF levels were attributed to decreased expression of liver genes, including Igf1, IGF binding, and ternary complex protein gene Igfals and Igfbp3. The previous studies suggest that the liver is the primary origin of systemic IGF, as demonstrated by the liver-specific deletion of Igf1 and the knockout of Igfals, which is mostly expressed in the liver (Yakar et al., 1999; Yakar et al., 2002). The double KO mice exhibited a 90% decrease in serum IGF levels and displayed a phenotype of shortened femur length and growth plates. It is thus possible that the decrease in serum IGF levels, resulting from reduced liver IGF pathway genes in SMA mice, has also played a role in the observed bone growth defect.

Mesenchymal progenitor cells (MPCs) derived from the lateral plate mesoderm (LPM) differentiate into various types of limb mesenchymal cells, including bone, cartilage, and intramuscular mesenchymal cells like fibro-adipogenic progenitors (FAPs) (Nassari et al., 2017). Recent studies have revealed the role of FAPs in skeletal muscle homeostasis, as reducing the number of FAPs resulted in diminished muscle regeneration capacity, long-term muscle atrophy, and NMJ denervation (Wosczyna et al., 2019; Uezumi et al., 2021). Our latest research demonstrated that FAP-specific deficiency of Bap1, one of the deubiquitinases, leads to NMJ defects (Kim et al., 2022). These recent findings raise the possibility that FAPs may have a specific role in the pathogenesis of neuromuscular diseases such as SMA. However, it has not been studied if depletion of SMN in FAPs can lead to SMA-like neuromuscular pathology.

In this study, we crossed a limb MPC-specific Cre mouse with a floxed Smn1 exon 7 mouse carrying multiple copies of the human SMN2 gene, allowing us to examine the impact of mesenchymal SMN reduction on SMA pathogenesis. As a result, the mutant mice showed skeletal growth abnormalities and local IGF signaling defects in the growth plate. In addition, our findings indicate that the SMN reduction in FAPs, similar to the extent of severe SMA, causes altered NMJ development.

Results

Bone growth restriction and growth plate defects caused by MPC-specific SMN depletion

To investigate the effects of SMN reduction within MPCs in SMA pathogenesis, we crossed Smn1f7/f7 mice, which possess loxP sites flanking exon 7 of the Smn1 gene (Frugier et al., 2000), with Prrx1Cre mice. This produced Smn1ΔMPC mice (Prrx1Cre; Smn1f7/f7) that lacked the Smn1 gene specifically in Prrx1Cre-expressed limb MPCs that give rise to bone, cartilage, and FAPs (Logan et al., 2002; Leinroth et al., 2022). To ascertain whether mutant mice carrying the SMN2 gene, like SMA patients, present pathological phenotypes, we additionally generated SMN2 2-copy Smn1ΔMPC (Prrx1Cre; Smn1f7/f7; SMN2+/+) and SMN2 1-copy Smn1ΔMPC (Prrx1Cre; Smn1f7/f7; SMN2+/0) mice. Control littermates that lacked Prrx1Cre were used as controls for comparison.

SMN2 0-copy Smn1ΔMPC mice died within 24 hr after birth. Regions where limbs should have formed at E18.5 only had rudimentary limb structures (Figure 1—figure supplement 1A). Furthermore, since the upper head bone did not cover the brain, it was directly attached to the skin and protruded. These observations can be attributed to Prrx1Cre-mediated SMN deletion in the LPM-derived limb MPCs and the craniofacial mesenchyme, which is accountable for the formation of calvarial bone (Wilk et al., 2017). To investigate whether the lack of SMN proteins in MPCs is responsible for bone development abnormalities, we performed alcian blue and alizarin red staining on E18.5 SMN2 0-copy Smn1ΔMPC mutants to analyze the structure of bones and cartilage. The appendages displayed restricted bone and cartilage formations, with scarcely discernible femur and tibia (Figure 1—figure supplement 1B, C). In the cranial region, there was an absence of both cartilage and bone at the location of the calvarial bone, with the parietal bone entirely missing and partially absent frontal bone (Figure 1—figure supplement 1D). Additionally, the sternum, which is one of the bones originating from the LPM (Sheng, 2015), was shorter than the control (Figure 1—figure supplement 1E).

The SMN2 2-copy Smn1ΔMPC mice, carrying two homologous SMN2 genes, did not show any discernible differences from the Prrx1Cre-negative control littermates into adulthood. However, the SMN2 1-copy Smn1ΔMPC mice exhibited reduced body size and shorter limb length compared to the SMN2 1-copy control (Smn1f7/f7; SMN2+/0). To assess postnatal bone growth defects observed in SMA patients and mouse models, we conducted micro-computed tomography (micro-CT) analysis on femurs obtained from postnatal day 14 (P14) SMN2 1-copy Smn1ΔMPC and SMN2 2-copy Smn1ΔMPC mice (Figure 1A). The 3D reconstruction image showed that the SMN2 1-copy mutant femur was smaller than the WT control and SMN2 2-copy mutant, and secondary ossification center is denied. The longitudinal virtual section view displayed reduced trabecular bone in the SMN2 1-copy mutant femur. CT analysis data showed that SMN2 1-copy mutants exhibited reduced femur diaphysis length, diameter, and trabecular bone volume compared to the control group, indicating growth plate-dependent endochondral ossification defects (Figure 1B–D). We then examined femoral bone thickness and diaphyseal bone mineral density (BMD) to determine whether mineralization was normal after bone formation. The thickness of the bone in the mutants did not differ significantly from the control, suggesting that bone mineralization was intact (Figure 1E and Figure 1—figure supplement 2A). Unexpectedly, BMD slightly increased in SMN2 1-copy mutants compared to the control group (Figure 1—figure supplement 2B). To assess the impact of osteoclasts and osteoblasts on diaphysis cortical bone mineralization, we utilized Itgb3 immunofluorescence as the osteoclast marker and toluidine blue staining for imaging bone-attached osteoblasts (Romeo et al., 2019; Colaianni et al., 2015). Osteoclast and osteoblast density did not significantly differ between the SMN2 1-copy mutant and the control (Figure 1—figure supplement 2C–E). The higher BMD may be attributed to greater mechanical stress caused by the shorter femur supporting the weight of the body, consistent with prior research indicating that elevated mechanical force leads to higher BMD in the femur (Hoxha et al., 2014; Ike et al., 2015). Nevertheless, the decrease in bone growth without apparent deterioration in bone mineralization of the femur of SMN2 1-copy Smn1ΔMPC mutants is consistent with findings from the whole-body SMA mouse model (Hensel et al., 2020). Collectively, these results suggest that mice carrying low copies of SMN2, with the Smn1 gene specifically deleted in MPCs, exhibit bone growth abnormalities, while osteoblast and osteoclast populations show no obvious defects based on our preliminary analyses.

Figure 1 with 2 supplements see all
Skeletal growth abnormalities and altered growth plate homeostasis in SMN2 1-copy Smn1ΔMPC mice.

(A) Representative 3D images and longitudinal section view of the ossified femur bone. Scale bars, 1 mm. (B, C) SMN2 1-copy mutant’s femurs showed reduced growth in diaphysis length and diameter, and (D) decreased trabecular bone volume. (E) Trabecular bone thicknesses were not significantly different between the control and mutant groups. The micro-computed tomography (micro-CT) analysis was performed in femur diaphysis and metaphysis from SMN2 1-copy Smn1ΔWT, SMN2 2-copy, and 1-copy Smn1ΔMPC mice at P14. One-way analysis of variance (ANOVA) with Tukey’s post hoc test, n = 3–5 mice in each genotype (B–E). (F) Representative images of hematoxylin and eosin (H&E) staining in the distal femur growth plate of control and mutant mice with 1 copy of SMN2 at P14. Scale bars, 100 μm. Resting zone (RZ), hypertrophic zone (HZ), and proliferative zone (PZ). (G–I) Indicated by black arrows, the HZ and PZ lengths were reduced in SMN2 1-copy Smn1ΔMPC mice, and the hypertrophic cell number in a section of the 1-copy mutant was decreased (n = 4 mice in each genotype; unpaired t-test with Welch’s correction). (J) Representative images of Ki67 immunostaining in the distal femur growth plate of control and mutant mice with 1 copy of SMN2 at P14. Scale bars, 100 μm, and (K) decreased Ki67+ percentage in resting zone chondrocytes. (L) Ki67+ percentage in the proliferative zone was not significantly different between the control and mutant groups. n = 3 mice in each genotype; unpaired t-test with Welch’s correction (K–L). ns: not significantly different. *p < 0.05; **p < 0.01. Error bars show standard error of the mean (SEM).

Various bone-forming cells originating from the LPM were known to contribute to bone formation, such as growth plate chondrocytes and osteoblasts. In SMN2 1-copy Smn1ΔMPC mutants, SMN may be deleted in these cells, suggesting that they play a role in the bone growth abnormalities observed in 2-week-old mice. Previous researchers revealed that primary osteoblasts from a severe SMA mouse model did not display notable differences from controls in an in vitro ossification test. And they did not observe any differences in bone voxel density and bone thickness in femurs at P3 severe SMA mice. This is supported by the absence of any bone thickness or BMD defects in the SMN2 1-copy mutant (Figure 1E and Figure 1—figure supplement 2A, B), and the unimpaired osteoblast population (Figure 1—figure supplement 2E). Thus, we conclude that the bone growth abnormalities observed in the 2-week-old SMN2 1-copy Smn1ΔMPC mutant are due to impaired endochondral ossification.

To determine whether bone growth defects in SMN2 1-copy Smn1ΔMPC mutants arise from disrupted chondrocyte homeostasis at growth plates, we stained the femur distal growth plate of P14 mice with hematoxylin and eosin (H&E; Figure 1F). In line with earlier findings in whole-body SMA mice (Hensel et al., 2020), SMN2 1-copy Smn1ΔMPC mice exhibited shorter proliferative and hypertrophic zones compared to control mice (Figure 1G, H). Additionally, there was a significant reduction in the number of chondrocytes in the hypertrophic zone (Figure 1I). To investigate the decrease in chondrocyte proliferation and subsequent reduction in the proliferative and hypertrophic zone, we stained the proliferation marker Ki67 in the growth plate of both SMN2 1-copy control and mutant samples (Figure 1J). We then quantified the percentage of Ki67+ nuclei in the resting and proliferative zones (Figure 1K, L). Although there was no significant difference observed in the Ki67+ percentage of proliferative zone chondrocytes in the proceeding proliferation state, there was an absolute reduction in resting zone chondrocyte proliferation. The decreased proliferation rate in the resting zone could have impeded the transition to the proliferative zone. Our data indicate that adequate expression of SMN is essential for the homeostasis of chondrocytes at growth plates.

Disruption of chondrocyte-derived IGF signaling in SMN2 1-copy Smn1ΔMPC mutants

The proliferation and differentiation of growth plate chondrocytes are regulated by systemic IGF (Shim, 2015; Karimian et al., 2011; Racine and Serrat, 2020). Previous research suggested that a key factor contributing to the pathological phenotype in SMA is the lowered expression of the Igf1/Igfbp3/Igfals genes, which produce IGF and IGF-carrying proteins, in the liver (Hua et al., 2011; Murdocca et al., 2012). As the IGF pathway proteins are downregulated in whole-body SMA mice, the bone growth defects observed in the mice have sparked debate as it remains unclear whether they are due to cell-autonomous defects by bone-forming cells’ SMN reduction, or the low liver-derived IGF level (Hua et al., 2011; Tsai et al., 2014; Deguise et al., 2021; Hensel et al., 2020).

To clarify this issue, we used Smn1ΔMPC mutant mice, which enabled us to investigate the effect of SMN depletion in bone-forming cells such as chondrocytes on bone growth, while circumventing the impact of the endocrine signal by Prrx1-negative organ. To investigate the impact of IGF signaling on growth plate chondrocytes, we employed immunofluorescence to evaluate the percentage of p-AKT-positive cells activated by the IGF–PI3K–AKT pathway in both SMN2 1-copy control and mutant femur distal growth plate (Figure 2A). Intriguingly, the percentage of p-AKT+ cells was significantly decreased in resting zone chondrocytes, but not in the proliferative zone, which aligns with the Ki67+ percentage (Figure 2B, C). Expectedly, the liver’s mRNA expression of IGF pathway genes, reported to be decreased in SMA mouse models and patients (Hua et al., 2011; Murdocca et al., 2012; Deguise et al., 2021; Sahashi et al., 2013), showed no difference when comparing controls to SMN2 1-copy Smn1ΔMPC mutants (Figure 2D). These findings indicate that the growth plate’s proliferation and hypertrophy in SMN2 1-copy mutants are affected by impairments in another AKT upstream signal rather than by liver-secreted systemic IGF.

Decreased chondrocyte-derived IGF–AKT axis by limb mesenchymal cell-specific survival motor neuron (SMN) depletion in SMN2 1-copy Smn1ΔMPC mice.

(A) Representative images of p-AKT immunostaining in distal femur growth plate from mice at P14. Scale bars, 100 μm. (B, C) The p-AKT-positive percentage was decreased in the resting zone chondrocytes, not in the proliferative zone (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). (D) Relative IGF axis mRNA expression in the livers of SMN2 1-copy control and mutant mice. The IGF pathway genes showed no difference when comparing controls to SMN2 1-copy Smn1ΔMPC mutants. (E, F) Relative IGF axis and chondrocyte differentiation marker mRNA expression in the chondrocytes of SMN2 1-copy control and mutant mice. The Igf1, Igfbp3, and hypertrophic marker Col10a1 expression were decreased in SMN2 1-copy Smn1ΔMPC mutants. n = 3 mice in each genotype; unpaired t-test with Welch’s correction (D–F). (G) Representative images of genomic PCR analysis from SMN2 1-copy Smn1ΔMPC mice tissues at P21. (H) Quantitative reverse transcription polymerase chain reaction (qRT-PCR) analysis from tissues of SMN2 1-copy Smn1WT and Smn1ΔMPC mice at P21 (n = 3 mice in each genotype; unpaired t-test with Welch’s correction). Deletion of Smn1 exon 7 was detected only in limb mesenchymal cells using genomic PCR (G) and full-length Smn1 mRNA expression (H). (I) Representative images of western blot analysis in cultured fibro-adipogenic progenitors (FAPs). SMN protein in FAPs of SMN2 1-copy Smn1ΔMPC mice exhibited a decrease comparable to that observed in the SMAΔ7 mice. (J) Relative SMN levels in cultured FAPs of the controls and mutants (n = 3 mice in each genotype; one-way analysis of variance (ANOVA) with Tukey’s post hoc test). ns: not significantly different. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.001. Error bars show standard error of the mean (SEM).

Figure 2—source data 1

Original file for the gel electrophoresis of genomic PCR in Figure 2G (Cre, Smn1F7, Smn1Δ7) and western blot analysis in Figure 2I (anti-alpha-tubulin, anti-SMN).

https://cdn.elifesciences.org/articles/92731/elife-92731-fig2-data1-v1.zip
Figure 2—source data 2

PDF containing Figure 2G, I and original scans of the PCR and western blot with highlighted bands and sample labels.

https://cdn.elifesciences.org/articles/92731/elife-92731-fig2-data2-v1.zip

There are reports indicating that local IGF expression plays a crucial role in bone development through the growth plate, in addition to circulating IGF (Hallett et al., 2019; Racine and Serrat, 2020). While most Igf1-null mice died before birth and had smaller tibial lengths than normal, liver-specific Igf1-deleted mice did not experience significant changes in body length or tibial length during postnatal growth, despite a 75% reduction in serum IGF-1 levels (Baker et al., 1993; Yakar et al., 1999). This indicates that local IGF in the growth plate is crucial for endochondral ossification, in addition to serum IGF. The chondrocyte-specific Igf1 knockout mouse demonstrated a reduction in postnatal body and femur length, and the chondrocyte-specific Igf1r knockout mouse demonstrated a significant reduction in bone growth, as well as a decrease in both growth plate proliferative and hypertrophic zone (Govoni et al., 2007; Wang et al., 2011). A recent study revealed that resting zone chondrocytes in the growth plate serve as a major source of local IGF and activate the p-AKT pathway via autocrine and paracrine IGF signaling (Oichi et al., 2023). The study further revealed that cells that constitute bone and bone marrow, apart from chondrocytes, do not express Igf1, making chondrocytes the solitary source of local IGF. These suggest that the growth plate defects and the reduction of resting zone AKT phosphorylation in SMN2 1-copy mutants may be due to chondrocyte-secreted IGF deficiency. To confirm this hypothesis, we evaluated the expression of IGF-related genes in chondrocytes from both SMN2 1-copy control and mutant femur (Figure 2E). Indeed, Igf1 and Igfbp3 were greatly depleted in SMN2 1-copy mutant chondrocytes. It is hypothesized that the increased presence of Ghr may be in response to the reduction of Igf1. As IGF directly causes chondrocyte hypertrophy (Wang et al., 1999), we assess the mRNA expression of chondrocyte hypertrophic marker Col10a1 and undifferentiated chondrocyte marker Col2a1 (Figure 2F). The results show that Col10a1 is decreased, while Col2a1 is increased in the mutant. In Figure 1H, I, the hypertrophic cell reduction may be caused by a low local IGF level. Therefore, depletion of local IGF in the growth plate of SMN2 1-copy mutants may hinder chondrocyte progression to proliferation and hypertrophy, leading to aberrations in endochondral ossification. In Figures 1L and 2C, it is possible that the serum IGF could affect the remnant proliferative and p-AKT-positive cells in the growth plate via the vascularization of bone marrow. Based on these findings, it was concluded that deprivation of SMN in chondrocytes leads to a decrease in local IGF signaling, which affects growth plate homeostasis.

Mesenchymal cell-specific SMN reduction similar to severe SMA mouse model in SMN2 1-copy Smn1ΔMPC mutants

To confirm the specific deletion of Smn1 in limb mesenchymal cells, including chondrocytes and FAPs, we performed quantitative reverse transcription polymerase chain reaction (qRT-PCR) for full-length Smn1 mRNA expressed by undeleted Smn1 allele. Our findings indicate that full-length Smn1 mRNA expression in the brain, liver, skeletal muscle, and spinal cord of SMN2 1-copy Smn1ΔMPC mice at postnatal day 21 (P21) was similar to that of control mice, while it was significantly reduced in isolated FAPs and chondrocytes (Figure 2H). Additionally, we confirmed the presence of the Smn1Δ7 variant, which is the exon 7-deleted form of Smn1 created by cre-lox-mediated recombination, in both FAPs and chondrocytes by conducting genomic PCR on the SMN2 1-copy Smn1ΔMPC mutant (Figure 2G). Since SMN was not downregulated in the tissues other than limb mesenchymal cells, non-mesenchymal cells were ruled out from being responsible for the phenotype observed in Smn1ΔMPC mutants.

Among our Smn1ΔMPC models, mice with two copies of SMN2 exhibit similar bone development parameters as control mice without SMN deletion. However, mice with one copy of SMN2 display bone pathological defects akin to SMA mouse models. This may be because the quantity of human full-length SMN protein produced by SMN2 2-copy, was sufficient to sustain SMN complex function in the SMN2 2-copy Smn1ΔMPC mutants, despite the absence of functional mouse SMN protein in limb mesenchymal cells. On the contrary, due to the lower expression of full-length SMN compared to SMN2 2-copy mutants, the SMN complex may not function properly in SMN2 1-copy mutant cells. We confirmed this by comparing the amount of full-length SMN protein in isolated FAPs from the hindlimbs of control, Smn1ΔMPC mutants, and a severe SMA mouse model (Smn1−/−; SMN2+/+; SMNΔ7+/+; SMAΔ7 mutants) (Figure 2I). The data show that SMN2 1-copy Smn1ΔMPC mutants exhibited ~80% reduction in SMN levels compared to the control group. The level of SMN protein in SMN2 1-copy Smn1ΔMPC mutants was similar to that in SMAΔ7 mutants. Conversely, SMN2 2-copy mutants display a decrease of approximately 40% in SMN protein levels compared to the control (Figure 2J). The moderately reduced expression of SMN is adequate to support regular bone development in SMN2 2-copy mutants. Taken together, our findings indicate that the reduction of mesenchymal SMN to levels comparable to that of the severe SMA mouse model causes SMA-like bone pathology in the SMN2 1-copy mutant.

Abnormal NMJ maturation in SMN2 1-copy Smn1ΔMPC mutants

To investigate whether disabling SMN in FAPs results in SMA-like neuromuscular impairments, we assessed if NMJ phenotypes observed in SMA mouse models also occur in SMN2 1-copy Smn1ΔMPC mutants. Both severe and mild SMA mouse models exhibit impaired NMJ maturation markers, including plaque-like morphology of acetylcholine receptor (AChR) clusters, neurofilament (NF) varicosities, and poor terminal arborization (Kong et al., 2009; Martinez et al., 2012; Monani et al., 2003; Kariya et al., 2008). To evaluate the impact of SMN deficiency in FAPs on NMJ maturation, we evaluated the NMJ maturation markers in the tibialis anterior (TA) muscles of control and SMN2 1-copy Smn1ΔMPC mutant mice at P21, a time when NMJ maturation is in progress (Figure 3A). Our examination revealed the presence of NF varicosities in SMN2 1-copy mutants as compared with control mice (Figure 3A, D). Additionally, the number of nerve branches was decreased and half of the total NMJs were poorly arborized in SMN2 1-copy mutants (Figure 3B, C). These presynaptic alterations are specific phenotypes in neurogenic atrophy-like SMA. Unlike neurogenic atrophy, physiologic atrophy shows no differences in presynaptic morphology, such as nerve branching (Deschenes et al., 2006). This suggests that the NMJ phenotypes observed in SMN2 1-copy Smn1ΔMPC mutant mice are not caused by decreased muscle size and activity resulting from bone growth abnormalities. The morphology of AChR clusters shows that the mutants have more immature plaque-like NMJs than the controls’ pretzel-like structure (Figure 3E). Therefore, these findings indicate that SMN2 1-copy Smn1ΔMPC mutants exhibit NMJ maturation abnormalities common in SMA mouse models.

Aberrant postnatal neuromuscular junction (NMJ) maturation in SMN2 1-copy Smn1ΔMPC mice.

(A) Immunostaining of NMJs in TA muscle of SMN2 1-copy Smn1WT and Smn1ΔMPC mice at P21 with anti-NF (green), anti-synaptophysin (blue), and α-Btx staining acetylcholine receptor (AChR; red). Scale bars, 20 μm. The confocal images of NMJs showed decreased presynaptic terminal branching and the existence of nerve terminal varicosities that were enlarged with neurofilament (NF; indicated by arrowheads) in the mutant. (B) The NMJs of the SMN2 1-copy mutant exhibited a significant decrease in presynaptic terminal arborization and (C) an increased percentage of poorly arborized NMJs (n = 3 mice in each genotype; unpaired t-test with Welch’s correction). (D) The percentage of NMJs exhibiting NF varicosities was higher in the SMN2 1-copy mutant group than in the control group (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). (E) For quantification of the NMJ maturation stage, we classified NMJs into five distinct developmental stages (Plaque: plaque-shaped endplate without any perforation; Small perforated: plaque-shaped endplate with small perforations; Large perforated: plaque-shaped endplate with large perforations; Open: C-shaped endplate; Pretzel: pretzel-like shaped endplate) and then compared the frequency patterns of SMN2 1-copy control and mutant mice (n = 3 mice in each genotype; two-way analysis of variance (ANOVA) with Tukey’s post hoc test). The NMJs of SMN2 1-copy mutants displayed plaque-like shapes, indicating that they were in the immature stage. (F) Immunostaining of NMJs in TA muscle of SMN2 1-copy Smn1WT and Smn1ΔMPC mice at P3 with anti-NF (green), anti-synaptophysin (blue), and α-Btx staining AChR (red). Scale bars, 10 μm. (G) There were no significant differences in AChR cluster size between the SMN2 1-copy control and mutant at P3 (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). (H) The ratio of the Synaptophysin area to the AChR area in NMJ was slightly higher in the SMN2 1-copy mutant at P3 (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). ns: not significantly different. *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001. All box-and-whisker plots show the median, interquartile range, minimum, and maximum. For the box-and-whisker plots, range bars show minimum and maximum (B, G, H). For the bar and line graph, error bars show standard error of the mean (SEM) (C–E).

Undisturbed NMJ formation in neonatal SMN2 1-copy Smn1ΔMPC mutants

To determine whether any NMJ defects were present prior to juvenile NMJ maturation in SMN2 1-copy Smn1ΔMPC, we examined NMJ formation in SMN2 1-copy Smn1ΔMPC mice at the neonatal stage on postnatal day 3. We evaluated the AChR and nerve terminal areas to assess post- and presynaptic development, respectively (Figure 3F). Measurements of AChR cluster size indicated no differences between control and SMN2 1-copy Smn1ΔMPC mice (Figure 3G). However, the area of AChR covered by nerve terminals was slightly larger in SMN2 1-copy Smn1ΔMPC (Figure 3H). We have no reasonable explanation for why the coverage is higher in the mutant. However, there does not appear to be abnormal development of the NMJ in the mutant, at least until the neonatal period. Therefore, we reasoned that SMN2 1-copy Smn1ΔMPC mutants began to exhibit deterioration in the NMJ maturation during the juvenile stage, following the intact neonatal development of NMJ.

Aberrant NMJ morphology in the adult SMN2 1-copy Smn1ΔMPC mice

To evaluate the organization of NMJ after the conclusion of postnatal NMJ development, considering mesenchymal SMN expression, we examined NMJ morphology in the TA muscle of control, SMN2 1-copy, and 2-copy Smn1ΔMPC mice at postnatal day 56 (P56). Our analysis revealed that presynapses were fragmented in SMN2 1-copy mutants, resulting in a bouton-like morphology, in contrast to the control and SMN2 2-copy Smn1ΔMPC mice (Figure 4A). In SMN2 1-copy Smn1ΔMPC mice, a twofold presynaptic fragmentation compared to the control was quantified, demonstrating nerve terminal shrinkage (Figure 4B). Additionally, NF ends displayed more severe varicosities than at P21 and were only connected to the proximal nerve by very thin NF, unlike control and SMN2 2-copy mice (Figure 4C). Remarkably, numerous presynaptic islands formed in SMN2 1-copy Smn1ΔMPC mice through the merging of fragmented presynapses and NF varicosity. In SMN2 1-copy mutants, AChR clusters displayed fragmented grape-shaped morphology that overlapped with nerve terminals, whereas control and SMN2 2-copy mice displayed pretzel-like structures (Figure 4D). These results suggest that defects in adult NMJ morphology occur when mesenchymal SMN protein is reduced to the extent of the SMN2 1-copy Smn1ΔMPC mutants.

Morphological deterioration in neuromuscular junctions (NMJs) of adult SMN2 1-copy Smn1ΔMPC mice.

(A) Immunostaining of NMJs in TA muscle of SMN2 1-copy Smn1WT, SMN2 2-copy, and SMN2 1-copy Smn1ΔMPC mice at P56 with anti-NF (green), anti-synaptophysin (blue), and α-Btx staining acetylcholine receptor (AChR; red). Scale bars, 20 μm. The confocal images of NMJs showed fragmentation and bouton-like neurofilament (NF) varicosities (indicated by arrowheads) in the SMN2 1-copy Smn1ΔMPC mice. The NMJs of the SMN2 1-copy mutant displayed fragmented presynapse (B), endplate (D), and NF varicosities (C) compared to SMN2 1-copy Smn1WT and SMN2 2-copy Smn1ΔMPC mice (n = 3–5 mice in each genotype; Presynaptic fragments and AChR fragments: Brown–Forsythe and Welch analysis of variance (ANOVA) with Games–Howell’s test; NF varicosities: one-way ANOVA with Tukey’s post hoc test). ns; not significantly different. ****p < 0.0001. All box-and-whisker plots show the median, interquartile range, minimum, and maximum. For the box-and-whisker plots, range bars show minimum and maximum (B, D). For the bar graph, error bars show standard error of the mean (SEM) (C).

Presynaptic neurotransmission alteration in SMN2 1-copy Smn1ΔMPC mutants

To investigate whether the morphologically aberrant NMJs of SMN2 1-copy Smn1ΔMPC mice have functional impairments, we isolated hindlimb extensor digitorum longus (EDL) muscles from P56 mice and conducted electrophysiological recording. We incubated the muscles with μ-conotoxin, which selectively inhibits muscle voltage-gated Na+ channels, prevented the induction of muscle action potential (Ling et al., 2010; Zanetti et al., 2018), and recorded the Miniature endplate potential (mEPP) and evoked endplate potential (eEPP) (Ling et al., 2010; Zanetti et al., 2018). mEPP, a response that occurs when spontaneously released acetylcholine binds to nicotinic AChR without nerve stimulation, was measured ex vivo near the NMJs of the EDL muscles in the control and SMN2 1-copy Smn1ΔMPC mice (Figure 5A). The mEPP amplitude was increased in SMN2 1-copy Smn1ΔMPC mice (Figure 5B), whereas mEPP frequency was comparable between the controls and mutants (Figure 5C). The results indicate that the NMJ synapses of SMN2 1-copy Smn1ΔMPC mice are functional and more sensitive to acetylcholine compared to the controls. Next, we measured the eEPP by stimulating an action potential at the peroneal nerve (Figure 5D). Despite the increased mEPP amplitude, the amplitude of eEPPs was significantly decreased in SMN2 1-copy Smn1ΔMPC mice (Figure 5E). These results suggest that the nerve terminals in SMN2 1-copy Smn1ΔMPC mice exhibit decreased quantal content. This could be due to a decrease in vesicle release probability. Notably, there was no difference in the paired-pulse response, indicating normal neurotransmitter release probability (Figure 5F, G). Taken together, these findings suggest that the presynaptic neurotransmission ability of the NMJ is reduced in SMN2 1-copy Smn1ΔMPC mutants.

Reduced presynaptic neurotransmission ability in the neuromuscular junctions (NMJs) of SMN2 1-copy Smn1ΔMPC mice.

(A) Representative traces of Miniature endplate potential (mEPP) from SMN2 1-copy Smn1ΔWT (top) and SMN2 1-copy Smn1ΔMPC (bottom) mice. (B, C) SMN2 1-copy mutant’s NMJs showed an increase in mEPP amplitude and no differences in mEPP frequency (1-copy control, n = 25, 9 mice; 1-copy mutant, n = 21, 8 mice; unpaired t-test with Welch’s correction). (D) Representative traces of evoked endplate potential (eEPP) from SMN2 1-copy Smn1ΔWT (top) and SMN2 1-copy Smn1ΔMPC (bottom) mice. (E) The mutant’s NMJs showed a stronger amplitude of eEPPs (1-copy control, n = 12, 4 mice; 1-copy mutant, n = 12, 3 mice; unpaired t-test with Welch’s correction). (F) Representative traces of paired-pulse response from SMN2 1-copy Smn1ΔWT (top) and SMN2 1-copy Smn1ΔMPC (bottom) mice. (G) Paired-pulse response was not different between SMN2 1-copy control and mutant NMJs, indicating a comparable neurotransmitter release probability (1-copy control, n = 8, 3 mice; 1-copy mutant, n = 6, 3 mice; unpaired t-test with Welch’s correction). The electrophysiological recording was performed in the extensor digitorum longus (EDL) muscle at P56. ns: not significantly different. *p < 0.05. All box-and-whisker plots show the median, interquartile range, minimum, and maximum. For the box-and-whisker plots, range bars show minimum and maximum (B, C, E, G).

Disturbed nerve terminal structure in SMN2 1-copy Smn1ΔMPC mice

To examine the NMJ ultrastructure of SMN2 1-copy Smn1ΔMPC mutants, we utilized transmission electron microscopy (TEM) (Figure 6A). The density of junctional folds in SMN2 1-copy Smn1ΔMPC mutant specimens was comparable to that of the control (Figure 6B). However, the density of synaptic vesicles was substantially elevated in the SMN2 1-copy mutants (Figure 6C). Since previous electrophysiological results suggested a decrease in presynaptic neurotransmission capacity in SMN2 1-copy mutants, this could be due to synaptic vesicles failing to fuse with the membrane, leading to the accumulation of vesicles in the terminal and reduced quantal contents in Figure 5. In Figure 3H, larger synaptophysin coverage in the mutant may be caused by this synaptic vesicle accumulation. Additionally, the detachment of the nerve terminal is more frequent at the NMJ of mutants (Figure 6D). The detachment of nerve terminals observed in SMN2 1-copy Smn1ΔMPC mutants could have also resulted in diminished presynaptic neurotransmission capacity. Collectively, these findings indicate that SMN2 1-copy Smn1ΔMPC mutants have nerve terminal-specific pathological defects at the NMJ ultrastructural level.

Abnormal nerve terminal ultrastructure in SMN2 1-copy mutant.

(A) Representative transmission electron microscopy (TEM) images from neuromuscular junctions (NMJs) of SMN2 1-copy Smn1ΔWT and SMN2 1-copy Smn1ΔMPC mice at P56. Scale bars, 500 nm. Nerve terminal (NT; indicated by the blue zone). Synaptic vesicles (SVs). Muscle fiber (MS; indicated by the red zone). Endplate junctional folds (JF). Nerve terminal detachment (indicated by arrow) was observed in SMN2 1-copy Smn1ΔMPC mice. (B) The density of junctional folds in the NMJ of SMN2 1-copy Smn1ΔMPC mice no significant change compared to the control, whereas (C) the density of synaptic vesicles was increased (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). (D) The detachment of the nerve terminal occurs more frequently at the NMJ of mutants (n = 3–4 mice in each genotype; unpaired t-test with Welch’s correction). ns: not significantly different. *p < 0.05; ***p < 0.001. All box-and-whisker plots show the median, interquartile range, minimum, and maximum. For the box-and-whisker plots, range bars show minimum and maximum (B, C). For the bar graph, error bars show standard error of the mean (SEM) (D).

FAPs transplantation rescues NMJ morphology in limb mesenchymal SMN mutants

SMN-deleted limb mesenchymal tissues in SMN2 1-copy Smn1ΔMPC mutants comprise not only FAPs, but also bone, cartilage, pericytes, and tendon, among others (Leinroth et al., 2022; Nassari et al., 2017). To evaluate the critical role of FAPs in the postnatal development of the NMJ, we isolated fluorescent protein-labeled wild-type FAPs from Prrx1Cre; Rosa26LSL-YFP/+ or Prrx1Cre; Rosa26 LSL-tdTomato/+ mice and transplanted them into the TA muscles of SMN2 1-copy Smn1ΔMPC mice at postnatal day 10. The TA muscle on the contralateral side was treated with a vehicle as a control. In SMN2 1-copy Smn1ΔMPC mice at P56, the muscle that received FAPs showed decreased presynaptic fragmentation, NF varicosities, and postsynaptic fragmentation compared to the contralateral muscle (Figure 7A–D). As a result, the transplanted muscle was not significantly different from the control except for NF varicosities. These data demonstrate that the transplantation of wild-type FAPs rescues the abnormal NMJ development in SMN2 1-copy Smn1ΔMPC mice. Overall, our findings indicate that SMN depletion in FAPs leads to the neuronal SMN-independent NMJ pathology in severe SMA, which is rescued by the transplantation of healthy FAPs.

Improved postnatal neuromuscular junction (NMJ) development in the TA muscle of SMN2 1-copy Smn1ΔMPC mice following healthy fibro-adipogenic progenitors (FAPs) transplantation.

(A) Immunostaining of NMJs in tdTomato+ FAP-transplanted TA muscle (+FAP) and vehicle-treated contralateral muscle (+Veh) in SMN2 1-copy Smn1ΔMPC mice at P56 with anti-NF (yellow), anti-synaptophysin (blue), α-Btx staining acetylcholine receptor (AChR; green), and tdTomato fluorescence (red). Scale bars, 40 μm. The images revealed neurofilament (NF) varicosities (indicated by arrowheads) in the +Veh NMJs. The tdTomato+ FAPs (marked by asterisks) were transplanted into +FAP NMJs, which exhibited (B) decreased presynaptic fragmentation, (C) NF varicosities, and (D) AChR fragmentation compared to +Veh NMJs and similar to wild-type NMJs (n = 3–4 mice in each group; unpaired t-test with Welch’s correction). ns: not significantly different. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. All box-and-whisker plots show the median, interquartile range, minimum, and maximum. For the box-and-whisker plots, range bars show minimum and maximum (B, D). For the bar graph, error bars show standard error of the mean (SEM) (C).

Discussion

In this paper, we elucidate the contribution of SMN depletion in mesenchymal progenitors for the pathogenesis of SMA. To test this hypothesis, we generated conditional knockout mouse strains to delete the Smn1 allele specifically in limb mesenchymal cells and carry human SMN2 copies. Our research using these mouse models resulted in three major discoveries. First, SMN deficiency in FAPs contributes to NMJ pathological defects in SMA. We observed delayed NMJ maturation and varicosities in juvenile SMN2 1-copy Smn1ΔMPC mutant. The pathogenic NMJ phenotypes were also observed in the SMAΔ7 mutant, which is one of the severe SMA mouse models (Kong et al., 2009; Martinez et al., 2012; Kariya et al., 2008). As the SMAΔ7 mutant typically lives for approximately 12 days, the fragmentation of the NMJ in adult SMN2 1-copy mutant was not evaluated in the severe SMA mutant. Nevertheless, models that induce a conditional adult SMN deficiency through either CreER allele or oligonucleotide administration resulting in SMN reduction, demonstrated the depletion of SMN throughout the body caused fragmentation of NMJ (Sahashi et al., 2013; Kariya et al., 2014). Thus, we demonstrate that the SMN2 1-copy Smn1ΔMPC mutant model mimics whole-body SMA mouse models in NMJ morphology. In the electrophysiological test, the SMAΔ7 mutant exhibited reduced quantal content, readily releasable pool, and vesicle release probability (Torres-Benito et al., 2011). In our study, we did not directly assess the readily releasable pool in SMN2 1-copy Smn1ΔMPC mutant by stimulus train of electrophysiological recording but instead showed reduced quantal content and normal vesicle release probability. In previous studies, it was theorized that the decrease in quantal contents in SMAΔ7 mutants resulted from decreased synaptic vesicle density at nerve terminals caused by motor neuron defects and abnormal axonal transport (Dale et al., 2011; Kong et al., 2009). However, we found that synaptic vesicle density was increased in the SMN2 1-copy Smn1ΔMPC mutant with SMN-sufficient motor neurons. It is possible that the alteration of active zones, which were also altered in the SMA motor terminals (Kong et al., 2009), contributed to the reduction in synaptic vesicle fusion and the decreased quantal contents. Indeed, nerve terminal detachment in SMN2 1-copy mutant mice was also found in active zone complex protein integrin-α3 knockout mice (Ross et al., 2017). Based on the rescue data of transplanted healthy FAPs, we can report that FAP-specific SMN depletion is involved in NMJ pathology of SMA.

Second, we demonstrated that skeletal growth defects, a phenotype observed in SMA (Khatri et al., 2008; Vai et al., 2015; Wasserman et al., 2017; Baranello et al., 2019; Hensel et al., 2020), are a cell-autonomous pathological effect of the depletion of SMN in bone-forming cells. We observed reduced bone size and volume in juvenile SMN2 1-copy Smn1ΔMPC mutant. Depletion of SMN in chondrocytes showed growth plate homeostasis problems with chondrocyte-secreted IGF defects. However, our skeletal study has a limitation in that we did not assess potential impairment of endochondral ossification during embryonic development. It may be necessary to further evaluate the embryonic period in addition to the postnatal growth plate defects observed in our study, as cells of the Prrx1-lineage are involved in mesenchymal condensation and chondrocyte differentiation during embryonic endochondral ossification (Hallett et al., 2019; Racine and Serrat, 2020). Previous studies have reported that IGF1 overexpression improves biochemical and behavioral manifestations in SMA mice, suggesting potential therapies for SMA (Bosch-Marcé et al., 2011; Tsai et al., 2014). Our study showed that these IGF therapies for SMA could be one to consider for treating bone growth abnormalities. In addition, the skeletal growth defect may affect physiologic muscle atrophy through imbalanced muscle contraction and reduced tension. This physiologic atrophy may be part of SMA-associated muscle weakness independent of neurogenic atrophy.

Third, it was demonstrated that adequate levels of SMN protein are essential for MPCs to contribute to limb neuromusculoskeletal development. The mutants with only 1 copy of SMN2 exhibit problematic symptoms observed in both SMA patients and mouse models, while SMN2 2-copy mutants display a typical phenotype in bone and NMJ. The lack of SMN protein in MPCs by insufficient SMN2 copies, similar to the deficiency seen in severe SMA, is responsible for the onset of SMA pathology. Thus, we propose that restoration of deficient SMN in MPCs is crucial for rehabilitating their function. Based on these discoveries, SMN replenishment treatments for MPCs, specifically FAPs, and chondrocytes, are necessary to provide a complete solution for neuromusculoskeletal defects in severe SMA patients.

The initial focus for the treatment of SMA was on motor neurons located in the spinal cord, and pharmaceuticals were created to address the deficiency of SMN in these neurons (Mercuri et al., 2020). For example, intrathecal injections of Spinraza can efficiently boost SMN in the CNS, including the spinal cord (Passini et al., 2011; Claborn et al., 2019). However, Spinraza does not address the lack of SMN in peripheral tissues, including mesenchymal cells, highlighted in recent studies. The drug Zolgensma, which employs AAV9 to express SMN through systemic delivery, appears to resolve this issue (Foust et al., 2009; Valori et al., 2010; Mattar et al., 2013). Nevertheless, a prior study indicates that chondrocytes within the growth plate and articular cartilage do not get infected with AAV9 (Yang et al., 2019). Furthermore, it is well established that DNA vectors delivered via AAV9 undergo dilution during cell proliferation (Penaud-Budloo et al., 2008; Colella et al., 2018; Van Alstyne et al., 2021; Heller et al., 2021). Given the infection issue and the dilution issue by the active cell population changes of chondrocytes and FAPs during postnatal development (Petrany et al., 2020; Bachman and Chakkalakal, 2022), SMN supplementation through Zolgensma alone would not be sufficient for severe SMA patients. Thus, there is an urgent need to research and develop therapeutic strategies that target mesenchymal progenitors.

While this study is the first to demonstrate the impact of SMN depletion in FAPs on the NMJ development, it does not elucidate the specific mechanism by which FAPs influence the NMJ. Our study observed NMJ recovery in the muscles transplanted with healthy FAPs, but not in the contralateral muscles, indicating that FAPs are likely involved in NMJ development through juxtacrine or paracrine signaling. Since FAPs interact with surrounding tissues through a variety of signaling factors, such as extracellular matrix (Contreras et al., 2021; Scott et al., 2019), Wnt-related protein (Lukjanenko et al., 2019), and Bmp signaling protein (Uezumi et al., 2021; Camps et al., 2020), mis-splicing of the signaling factors due to SMN reduction could disturb the homeostasis of neighboring tissues (Zhang et al., 2008). Furthermore, the Hsd11b1-positive subpopulation of FAPs associated with NMJ was discovered through single-cell RNA sequencing in a recent study (Leinroth et al., 2022). This population is located adjacent to the NMJ and responds to denervation, indicating an increased possibility of interaction with the NMJ organization. Therefore, it is necessary to conduct additional investigations into the expression of various signaling factors by diverse FAP subpopulations in future studies.

Methods

Key resources table
Reagent type (species) or resourceDesignationSource or referenceIdentifiersAdditional information
Genetic reagent (Mus musculus)Prrx1CreThe Jackson LaboratoryStrain #: 005584;
RRID:IMSR_JAX:005584
Genetic reagent (M. musculus)Smn1f7/+The Jackson LaboratoryStrain #: 006138; RRID:IMSR_JAX:006138
Genetic reagent (M. musculus)Rosa26LSL-YFP/+The Jackson LaboratoryStrain #: 006148; RRID:IMSR_JAX:006148
Genetic reagent (M. musculus)Rosa26LSL-tdTomato/+The Jackson LaboratoryStrain #: 007914; RRID:IMSR_JAX:007914
Genetic reagent (M. musculus)SMN2+/+; Smn1+/−The Jackson LaboratoryStrain #: 005024; RRID:IMSR_JAX:005024
Genetic reagent (M. musculus)Smn1+/−; SMN2+/+; SMNΔ7+/+The Jackson LaboratoryStrain #: 005025; RRID:IMSR_JAX:005025
Antibodyanti-SMN (Mouse monoclonal)BD BiosciencesCat. #: 610646;
RRID:AB_397973
WB (1:1000)
Antibodyanti-alpha-tubulin (Rabbit monoclonal)AbcamCat. #: ab176560; RRID:AB_2860019WB (1:1000)
Antibodyanti-Neurofilament M (Rabbit polyclonal)MerckmilliporeCat. #: ab1987; RRID:AB_91201IF (1:1000)
Antibodyanti-Synaptophysin 1 (Guinea pig polyclonal)Synaptic SystemsCat. #: 101 004; RRID:AB_1210382IF (1:500)
Antibodyanti-GFP (Chicken polyclonal)AbcamCat. #: ab13970; RRID:AB_300798IF (1:500)
Antibodyanti-Ki67 (Rabbit polyclonal)AbcamCat. #: ab15580; RRID:AB_443209IF (1:500)
Antibodyanti-Itgb3 (Rabbit monoclonal)Cell SignalingCat. #: 13166; RRID:AB_2798136IF (1:100)
Antibodyanti-p-AKT(S473) (Rabbit monoclonal)Cell SignalingCat. #: 4060; RRID:AB_2315049IF (1:100)
Antibodyanti-CD45-APC (Rat monoclonal)BiolegendCat. #: 103111; RRID:AB_312976FACS (3 µl per test)
Antibodyanti-CD31-APC (Rat monoclonal)BiolegendCat. #: 102409; RRID:AB_312904FACS (3 µl per test)
Antibodyanti-Sca-1(Ly6a)-FITC (Rat monoclonal)BiolegendCat. #: 122507; RRID:AB_756192FACS (3 µl per test)
Antibodyanti-Sca-1(Ly6a)-Pacific blue (Rat monoclonal)BiolegendCat. #: 108120; RRID:AB_493273FACS (3 µl per test)
Antibodyanti-Vcam1-Biotin (Rat monoclonal)BiolegendCat. #: 105703; RRID:AB_313204FACS (3 µl per test)
Antibodyanti-Rabbit IgG-HRP (Goat monoclonal)PromegaCat. #: W4011; RRID:AB_430833WB (1:10,000)
Antibodyanti-Mouse IgG-HRP (Goat monoclonal)PromegaCat. #: W4021; RRID:AB_430834WB (1:10,000)
Antibodyanti-Rabbit IgG-Alexa fluor 488 (Goat monoclonal)InvitrogenCat. #: A11034; RRID:AB_2576217IF (1:500)
Antibodyanti-Chicken IgY-Alexa fluor 488 (Goat monoclonal)InvitrogenCat. #: A11039; RRID:AB_2534096IF (1:500)
Antibodyanti-Rabbit IgG-Alexa fluor Plus 647 (Goat monoclonal)InvitrogenCat. #: A32733; RRID:AB_2633282IF (1:500)
Antibodyanti-Guine pig IgG-Alexa fluor 405 (Goat monoclonal)AbcamCat. #: ab175678; RRID:AB_2827755IF (1:500)
Peptide, recombinant proteinPE-Cy7-StreptavidinBiolegendCat. #: 405206FACS (3 µl per test)
Peptide, recombinant proteinAlpha-bungarotoxin-Alexa fluor 555InvitrogenCat. #: B35451IF (1:1000)
Peptide, recombinant proteinAlpha-bungarotoxin-Alexa fluor 488InvitrogenCat. #: B13422IF (1:1000)
Peptide, recombinant proteinμ-conotoxin GIIIBAlomoneCat. #: C-270Electrophysiology (2.5 µM)
Commercial assay or kitPierce BCA protein assay kitsThermo
Fisher
Scientific
Cat. #: C-23225
SoftwareImageJImageJRRID:SCR_003070
SoftwareAccuCTPerkinElmer
SoftwareMicrosoft ExcelMicrosoftRRID:SCR_016137
SoftwareLeica Application Suite XLeicaRRID:SCR_013673
SoftwareGraphPad PrismGraphpadRRID:SCR_002798
SoftwarepClampMolecular devicesRRID:SCR_011323
SoftwareQIAGENQIAGENRRID:SCR_008539
OtherGill No. 3 formula hematoxylinSigmaCat. #: GHS332Hematoxylin staining solution (1×)
OtherEosin Y solutionSigmaCat. #: HT110116Eosin staining solution (1×)
OtherToluidine blueSigmaCat. #: 198161Toluidine blue staining (1%)

Animals

Prrx1Cre (stock 005584), Smn1f7/+ (stock 006138), Rosa26 LSL-YFP/+ (stock 006148), Rosa26 LSL-tdTomato/+ (stock 007914), SMN2+/+, Smn1+/− (stock 005024 – Smn1 knockout and SMN2 homologous transgenic mouse) and Smn1+/−; SMN2+/+; SMNΔ7+/+ (stock 005025 005024 – Smn1 knockout and SMN2, SMNΔ7 homologous transgenic mouse) mice were acquired from The Jackson Laboratory (Bar Harbor, ME, USA). SMN2 0-copy Smn1ΔMPC mice (Prrx1Cre; Smn1f7/f7), SMN2 1-copy Smn1ΔMPC mice (Prrx1Cre; Smn1f7/f7; SMN2+/0 – SMN2 heterologous allele), and SMN2 2-copy Smn1ΔMPC mice (Prrx1Cre; Smn1f7/f7; SMN2+/+ – SMN2 homologous allele) were generated by crossing Prrx1Cre mice with Smn1f7/+ and Smn1+/−; SMN2+/+mice. To utilize FAPs transplantation, we generated Prrx1Cre; Rosa26 LSL-YFP/+ mice and Prrx1Cre; Rosa26 LSL-tdTomato/+ mice by breeding Prrx1Cre with Rosa26 LSL-YFP/+ and Rosa26 LSL-tdTomato/+ mice, respectively. To avoid deletion of the floxed allele by Prrx1Cre expression in female germline, we bred females without Prrx1Cre line to Prrx1Cre transgenic males. SMAΔ7 mutants (Smn1−/−; SMN2+/+; SMNΔ7+/+) were produced by mating Smn1+/−; SMN2+/+; SMNΔ7+/+ mice. Both male and female mice were used in the experiments, and no sex-specific differences were observed. Control littermates lacking Prrx1Cre were utilized for analysis. All mouse lines were housed under controlled conditions with specific pathogen free and handled according to the guidelines of the Seoul National University Institutional Animal Care and Use Committee (Protocol number: SNU-210313-1).

Micro-CT

The femurs from three groups – control, SMN2 2-copy Smn1ΔMPC, and SMN2 1-copy Smn1ΔMPC mice at postnatal day 14 (P14) – were isolated and cleaned of muscles and skin. Subsequently, the femurs were preserved in 4% paraformaldehyde (PFA) in phosphate-buffered saline (PBS) overnight at 4°C before micro-CT. The femurs were then imaged through a Quantum GX II micro-CT imaging system (PerkinElmer, Hopkinton, MA, USA). The X-ray source for scanning was set at 90 kV and 88 mA with a field of view of 10 mm (voxel size, 20 μm; scanning time, 14 min). The 3D imaging was viewed using the 3D Viewer software of the Quantum GX II. The size and volume of the femur bone were measured via AccuCT analysis software within the ossified diaphysis and metaphysis of the femur, excluding the epiphysis. BMD calibration was performed using a 4.5-mm BMD phantom and BMD measurements were taken at the center of the diaphysis.

Histology

Alcian blue and alizarin red staining was performed on control and SMN2 0-copy Smn1ΔMPC mice at E18.5 using a previously reported protocol (Ovchinnikov, 2009). For H&E and toluidine blue staining of the growth plate in the control and SMN2 1-copy Smn1ΔMPC mice at P14, the femurs were fixed in 4% PFA for 24 hr, rinsed in running tap water for 24 hr, and incubated with 10% ethylenediaminetetraacetic acid (EDTA) (pH 7.4) at 4°C with shaking for 2–3 days. Subsequently, the samples were rinsed in running tap water for 24 hr and then dehydrated through ethanol/xylene and embedded in paraffin. The embedded samples were then sectioned to a thickness of 5 µm, rehydrated, and stained with Gill No. 3 formula hematoxylin and eosin Y (H&E, Sigma-Aldrich, St. Louis, MO, USA) and toluidine blue (Sigma-Aldrich, St. Louis, MO, USA). Stained slides were analyzed using ×10 and ×20 objectives in the EVOS M7000 imaging system (Thermo Fisher Scientific, Waltham, MA, USA). The growth plate’s proliferative and hypertrophic zones were defined by their respective cell sizes.

Hindlimb FAPs isolation

Isolation of limb muscle FAPs was performed according to a previously reported protocol (Kim et al., 2020) with modifications. Limb muscles were dissected and mechanically dissociated in Dulbecco’s modified Eagle’s medium (DMEM, Hyclone) containing 10% horse serum (Hyclone, Logan, UT, USA), collagenase II (800 units/ml; Worthington, Lakewood, NJ, USA), and dispase (1.1 units/ml; Thermo Fisher Scientific, Waltham, MA, USA) at 37°C for 40 min. Digested suspensions were subsequently triturated by sterilized syringes with 20 G 1/2 needle (BD Biosciences, Franklin Lakes, NJ, USA) and washed with DMEM to harvest mononuclear cells. Mononuclear cells were stained with corresponding antibodies. All antibodies used in fluorescence-activated cell sorting (FACS) analysis are listed in Supplementary file 1. To exclude dead cells, 7AAD (Sigma-Aldrich; St. Louis, MO, USA) was used. Stained cells were analyzed and 7AADLinVcamSca1+ (stem cell antigen 1; Ly6a) (FAPs) were isolated using FACS Aria III cell sorter (BD Biosciences) with four-way purity precision. For western blot, freshly isolated FAPs were cultured at 37°C in alpha-MEM (Hyclone) supplemented with Antibiotic–Antimycotic (anti–anti, Gibco) and 20% fetal bovine serum (FBS; Hyclone). For transplantation, YFP+ FAPs and tdTomato+ FAPs were isolated from postnatal day 10–21 Prrx1Cre; Rosa26 LSL-YFP/+ mice and Prrx1Cre; Rosa26 LSL-tdTomato/+ mice, respectively. Isotype control density plots were used as a reference for positive gating.

Chondrocytes isolation

Isolation of chondrocytes followed a modified protocol previously published (Jonason et al., 2015). Using blunt forceps, cartilage caps were removed from P21 mouse femoral heads and dissected into ~1 mm fragments in a Petri dish with 10× anti–anti in PBS. The cartilage fragments were washed twice with PBS and then incubated in 5 ml collagenase II solution (800 units/mL collagenase II in DMEM with anti–anti, sterilized by 0.2 µm filtration) in a 60 mm culture dish at 37°C in a 5% CO2 incubator overnight. Chondrocytes were released by pipetting the remaining cartilage fragments 10 times and then filtering them through a 70-μm cell strainer to a 50-ml conical tube. The cells were then washed twice with PBS and pelleted by centrifugation at 500 × g for 5 min. Subsequently, the cells were cultured overnight at 37°C in a 5% CO2 incubator, in complete culture medium (DMEM with 10% FBS and anti–anti) overnight at 37°C in a 5% CO2 incubator.

PCR reaction

To detect the presence of Smn1 exon 7 floxed and deleted allele, 20 mg of each tissue were dissolved in direct PCR buffer (VIAGEN) with proteinase K overnight at 65°C. After inactivation at 95°C for 30 min, PCR was performed using previously reported primer sets to verify the existence of Cre, Smn1f7, and Smn1Δ7 alleles (Frugier et al., 2000). Primers are listed in .

RNA extraction and measurement of mRNA expression

Total RNA was extracted from the brain, liver, spinal cord L4, tibialis anterior muscle, freshly isolated FAPs, and chondrocytes using a TRIzol Reagent (Life Technologies, Carlsbad, CA, USA) and analyzed by qRT-PCR. First-strand complementary DNA was synthesized from 1 μg of RNA using ReverTra Ace (Toyobo, Osaka, Japan) containing random oligomer according to the manufacturer’s instructions. qRT-PCR (QIAGEN) was performed with SYBR Green technology (SYBR Premix Ex Taq, QIAGEN) using specific primers against indicated genes. Relative mRNA levels were determined using the 2−ΔΔCt method and normalized to Gapdh (Figure 1H, Figure 1—figure supplement 1K). Primers are listed in Supplementary file 1.

Western blot

Cultured FAPs at passage 3 were homogenized in radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris–HCl, pH 7.5, 0.5% sodium dodecyl sulfate, 20 µg/ml aprotinin, 20 µg/ml leupeptin, 10 µg/ml phenylmethylsulfonyl fluoride, 1 mM sodium orthovanadate, 10 mM sodium pyrophosphate, 10 mM sodium fluoride, and 1 mM dithiothreitol). Cell lysates were centrifuged at 13,000 rpm for 15 min. Supernatants were collected and subjected to immunoblot. BCA protein assay (Thermo Fisher Scientific) was used for estimating total protein concentrations. Normalized total proteins were analyzed by electrophoresis in 10% polyacrylamide gels and transferred to polyvinylidene fluoride (PVDF) membranes (Millipore, Billerica, MA, USA). Membranes were blocked in 5% skim milk (BD Biosciences) in tris-buffered saline (TBS) with 0.1% Tween-20 and incubated with primary antibodies overnight at 4°C. After incubation with the corresponding horseradish peroxidase (HRP)-conjugated secondary antibodies, the membranes were developed using a Fusion solo chemiluminescence imaging system (Vilber, Marne-la-Vallée, France). α-Tubulin was used as a loading control. Band intensities were quantified using ImageJ software. Antibodies used in this study are listed in Supplementary file 1. Primary and secondary antibodies were diluted 1:1000 and 1:10,000 with PBS containing 0.1% Tween-20 and 3% bovine serum albumin, respectively.

FAPs transplantation

FAPs transplantation was performed according to a previously reported protocol (Kim et al., 2020) with modifications. YFP+ or tdTomato+ FAPs (7AADLinVcamSca1+) were isolated by FACS from the limb muscles of indicated mice. 1 × 105 FAPs were suspended in 0.1% gelatin (Sigma-Aldrich, St. Louis, MO, USA) in PBS and then transplanted into one side of the TA muscles of SMN2 1-copy Smn1ΔMPC mice. The contralateral muscle received an equivalent volume of 0.1% gelatin in PBS (Vehicle).

Electrophysiology

The EDL muscle was dissected from control and SMN2 1-copy Smn1ΔMPC mice, along with the peroneal nerve, and then pinned to a Sylgard-coated recording chamber. Intracellular recording was conducted in oxygenated Ringer’s solution, which comprised 138.8 mM NaCl, 4 mM KCl, 12 mM NaHCO3, 1 mM KH2PO4, 1 mM MgCl2, and 2 mM CaCl2 with a pH of 7.4. Action potential of the muscle was prevented by preparing the muscle in 2.5 μM μ-conotoxin GIIIB (Alomone, Jerusalem, Israel) for 10 min beforehand. The recording was performed in toxin-free Ringer’s solution. mEPPs were recorded from a junction, followed by recordings of eEPPs by stimulating the attached peroneal nerve. The eEPPs were elicited using evoked stimulation. Paired-pulse stimulation (10-ms interstimulus interval) was utilized to assess synaptic transmission. The data were obtained and analyzed with Axoclamp 900A and Clampfit version 10.7 software.

Transmission electron microscopy

NMJ TEM followed a modified protocol previously reported (Modla et al., 2010). Mouse EDL muscle was swiftly excised and fixed in 4% PFA dissolved in Sorensen’s phosphate buffer (0.1 M, pH 7.2), followed by washing in 0.1 M phosphate buffer. The EDL was then gradually infiltrated on a rotator at room temperature with sucrose: 0.1 M phosphate buffer solutions of 30% and 50%, for 1 hr each, followed by an overnight incubation in 70% sucrose. Excess sucrose was then eliminated using filter paper, and the muscle was embedded in an optimal cutting temperature compound (O.C.T.; Sakura Fineteck, Torrance, CA, USA), followed by being frozen in a cryostat (Leica, Wetzlar, Germany). 10-µm-thick longitudinal sections were washed in PBS and treated with Alexa fluor 555-conjugated α-bungarotoxin (1:500, Invitrogen) for an hour. Imaging was conducted with the EVOS M7000 imaging system, and we selected four to five NMJ-rich regions for processing with TEM. The sections were fixed with 2% glutaraldehyde and 2% PFA in 0.1 M cacodylate buffer (pH 7.2) for 2 hr at room temperature, with an additional overnight incubation at 4°C. After washing with 0.1 M cacodylate buffer they were post-fixed with 1% osmium tetraoxide in 0.1 M cacodylate buffer (pH 7.2) for 2 hr at 4°C. The sections were then stained en bloc with 0.5% uranyl acetate overnight, washed with distilled water, and dehydrated using serial ethanol and propylene oxide. The sections were embedded in epoxy resin (Embed-812, Electron Microscopy Sciences) and detached from the slides by dipping them in liquid nitrogen. Ultra-thin sections (70 nm) were prepared with a diamond knife on an ultramicrotome (ULTRACUT UC7, Leica) and mounted on 100 mesh copper grids. Sections were stained with 2% uranyl acetate for 10 min and lead citrate for 3 min, then observed using a transmission electron microscope (80 kV, JEM1010, JEOL or 120 kV, Talos L120C, FEI). Synaptic vesicle density was quantified within a distance of 500 nm from the presynaptic membrane.

Immunohistochemistry

For NMJ staining, freshly dissected TA muscles were fixed in 4% PFA for 30 min at room temperature. Subsequently, the muscles were cryoprotected in 30% sucrose overnight, embedded in O.C.T., snap-frozen in liquid nitrogen, and stored at −80°C prior to sectioning. Longitudinal 40-μm-thick sections were obtained from the embedded muscles using a cryostat. The sections were blocked for 2 hr at room temperature using 5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100. Then, the sections were incubated with primary antibodies in the blocking buffer for 2 days at 4°C. After washing the sections three times with PBS/0.4% Triton X-100, the sections were stained with secondary antibodies overnight at 4°C, and then incubated with Alexa fluor 488- or 555-conjugated α-bungarotoxin (1:500, Invitrogen) for 2 hr at room temperature (RT), washed three times with PBS/0.4% Triton X-100 and mounted in VECTASHIELD. Z-serial images were collected at ×40 with a Leica SP8 confocal laser scanning microscope. To analyze NMJ morphology, LasX software was used to obtain maximal projections. NF varicosity refers to the varicose NF end connected to the rest of the nerve terminal. To quantify NMJ size and synaptophysin coverage, the Btx area and synaptophysin area were measured by ImageJ analysis software.

For bone section staining, the 5-µm-thick bone sections were rehydrated and antigen retrieval was then performed in citrate buffer (10 mM citric acid, pH 6) at 95°C 20 min. The sections were blocked for 1 hr at room temperature using 5% goat serum and 5% bovine serum albumin in PBS/0.4% Triton X-100. Then, the sections were incubated with primary antibodies in the blocking buffer at 4°C overnight. After washing the sections three times with PBS/0.1% Triton X-100, the sections were stained with secondary antibodies for 1 hr at RT, washed and mounted. Imaging was conducted with the EVOS M7000 imaging system.

Statistical analysis

Sample size determination was based on anticipated variability and effect size that was observed in the investigator’s lab for similar experiments. For quantification, individual performing the counts were blinded to sample identity and randomized. All statistical analyses were performed using GraphPad Prism 9 (GraphPad Software). For comparison of significant differences in multiple groups for normally distributed data, statistical analysis was performed by one- or two-way analysis of variance (ANOVA) followed by Tukey’s pairwise comparison post hoc test. For non-normally distributed data, Brown–Forsythe and Welch ANOVA followed by Games–Howell multiple comparisons test was used. For the comparison of two groups, Student’s unpaired t-test assuming a two-tailed distribution with Welch’s correction was used. Unless otherwise noted, all error bars represent standard error of the mean. The number of biological replicates and statistical analyses for each experiment were indicated in the figure legends. Independent experiments were performed at least in triplicates. p < 0.05 was considered statistically significant at the 95% confidence level. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001.

Data availability

All data generated or analyzed during this study are included in the manuscript and supporting files; source data files have been provided for Figure 2.

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Peer review

Reviewer #1 (Public Review):

Summary:

The manuscript by Hann et. al examines the role of survival motor neuron protein (SMN) in lateral plate mesoderm-derived cells using the Prrx1Cre to elucidate how changing cell-specific SMN levels coordinate aspects of the spinal muscular atrophy (SMA) pathology. SMN has generally been studied in neuronal cells, and this is one of the first insights into non-neuronal cells that may contribute to SMA disease. The authors generated 3 mouse lines: a Prrx1;Smnf/f conditional null mouse, as well as, single and double copy Prrx1;Smnf/f;SMN2 mice carrying either one or two copies of a human SMN2 transgene. First, the bone development and growth of all three were assessed; the conditional null Smn mutation was lethal shortly after birth, while the SMN2 2-copy mutant did not exhibit bone growth phenotypes. Meanwhile, single-copy SMN2 mutant mice showed reduced size and shorter limbs with shorter proliferative and hypertrophic chondrocyte zones. The authors suggested that this was cell autonomous by assessing the expression of extrinsic factors known to modulate proliferation/differentiation of growth plate chondrocytes. After assessing bone phenotypes, the authors transitioned to the assessments of neuromuscular junction (NMJ) phenotypes, since there are documented neuromuscular impairments in SMA and the Prrx1Cre transgene is expressed in muscle-associated fibro-adipogenic progenitors (FAPs). Neonatal NMJ development was unchanged in mutant mice with two copies of SMN2 , but adult single-copy SMN2 mutant mice had abnormal NMJ morphology, altered presynaptic neurotransmission, and problematic nerve terminal structure. Finally, the authors sought to assess the ability to rescue NMJ phenotypes via FAP cell transplantation and showed wild-type FAPs were able to reduce pre/postsynaptic fragmentation and neurofilament varicosities.

Strengths:

The conditional genetic approaches are novel and interestingly demonstrate the potential for chondrocyte and fibro-adipogenic progenitor-specific contributions to the SMA pathology.

The characterizations of the neuromuscular and NMJ phenotypes are relatively strong.

The data strongly suggest a non-neuronal contribution to SMA, which indicates a need for further mechanistic (cellular and molecular) studies to better understand SMA.

Weaknesses:

The skeletal analyses are not rigorous and likely do not get to the core of how SMN regulates bone development.

The overall work is descriptive and lacks convincing mechanisms.

Additional experimentation is likely needed to fully justify the conclusions.

https://doi.org/10.7554/eLife.92731.3.sa1

Reviewer #2 (Public Review):

Summary:

Sang-Hyeon et al. laid out a compelling rationale to explore the role of the SMN protein in mesenchymal cells, to determine whether SMN deficiency there could be a pathologic mechanism of SMA. They crossed Smnf7/f7 mice with Prrx1Cre mice to produce SmnΔMPC mice where exon 7 was specifically deleted and thus SMN protein was eliminated in limb mesenchymal progenitor cells (MPCs). To demonstrate gene dosage-dependence of phenotypes, SmnΔMPC mice were crossed with transgenic mice expressing human SMN2 to produce SmnΔMPC mice with different copies of SMN2 (0, 1, or 2). The paper provides genetic evidence that SMN in mesenchymal cells regulates the development of bones and neuromuscular junctions. Genetic data were convincing and revealed novel functions of SMN.

Strengths:

Overall, the paper provided genetic evidence that SMN deficiency in mesenchymal cells caused abnormalities in bones and NMJs, revealing novel functions of SMN and leading to future experiments. As far as genetics is concerned, the data were convincing (except for the rescue experiment, see below); the conclusions are important.

Weaknesses:

The paper seemed to be descriptive in nature and could be improved with more experiments to investigate underlying mechanisms. In addition, some data appeared to be contradicting or difficult to explain. The rescue data were not convincing.

https://doi.org/10.7554/eLife.92731.3.sa2

Reviewer #3 (Public Review):

Summary:

SMN expression in non-neuronal cells, particularly in limb mesenchymal progenitors is essential for the proper growth of chondrocytes and the formation of adult NMJ junctions.

Strengths:

The authors show copy numbers of smndelta7 in MPC influence NMJ structure.

Weaknesses:

Functional recovery by FAP transplantation is not complete. Mesenchymal progenitors are heterogeneous, and how heterogeneity influences this study is not clear. Part of the main findings to show the importance of SMN expression in non-neuronal cells is partly published by the same group (Kim et al., JCI Insight 2022). In the study, the authors used Dpp4(+) cells. The difference between the current study and the previous study is not so clear.

https://doi.org/10.7554/eLife.92731.3.sa3

Author response

The following is the authors’ response to the original reviews.

Reviewer #1

Major comments:

1. The authors conclude that the bone growth defects are chondrocyte-specific, highlighting no changes in the IGF pathway. However, other bone cells such as mesenchymal progenitors, osteoblasts, osteocytes, and marrow stromal cells are also lateral plate mesoderm derived and likely have roles in the bone growth phenotypes (a). Additionally, while the size decrease of the proliferative zone was stated, no actual proliferation assays such as BrdU were conducted (b). With the elements being of such small size in the mutants, the defects are likely to be found at the earliest stages of limb development at E11.5-E13.5 and may be due to mesenchymal to chondrocyte transitions or defects in osteoblast lineage development (c). Overall, the skeletal characterization is not rigorous and does not identify even a likely cellular mechanism. Further, a molecular mechanism by which SMN functions in mesenchymal progenitors, chondrocytes, or osteoblast lineage cells has not been assessed (d).

(a, c) As the reviewer commented, it seems to be a very important point to evaluate whether there is any problem in embryonic development from the time of mesenchymal cell condensation of the limb bud to the primary ossification center. However, when Hensel et al evaluated bone growth in P3 of severe SMA mice, the growth defect was not very large, with control femur length 3.5 mm and mutant 3.2 mm. it seems that even if SMN defects occur, there is no major problem with endochondral bone formation in the embryonic period (Hensel et al., 2020).

In this study, the SMN2 1-copy mutant with the bone growth defect was found to have a similar reduction in SMN protein to the severe SMA mouse model in experiments quantifying SMN protein. When Hensel et al. performed an in vitro ossification test on primary osteoblasts from the other severe SMA mouse model (Taiwanese severe SMA), they found no significant difference compared to controls. In femurs at P3 from severe SMA mice, they found no difference in bone voxel density and bone thickness (Hensel et al., 2020). In our data, bone thickness was not different in Figure 1 and Figure 1 – figure supplement 2, and BMD was actually greater. Thus, we believe that osteoblast and osteocyte function does not appear to be impaired by the absence of SMNs. When we looked at cortical osteoblasts in our new Figure 1-figure supplement 2, there did not appear to be a significant difference in density.

Furthermore, it is unlikely that BMSCs contributed to the bone growth we observed up to 2 weeks of age. the Lepr+Cxcl12+ BMSC population, which constitutes 94% ± 4% of CFU-F colonies formed by bone marrow cells (Zhou et al.k, 2014), is Prrx1-positive, and is known to be capable of osteogenesis in vivo, was only shown to differentiate into osteoblasts and form new bone in adults over 8 weeks of age. In the Lepr-cre; tdTomato; Col2.3-GFP mouse model, few cells expressing the osteoblast marker Col2.3-GFP are found before 2 months, and only about 3% of femur trabecular and cortical osteocytes express tdTomato at 2 months (Zhou et al., 2014). In Cxcl12-CreER; tdTomato; Col2.3-GFP mouse model, the researchers did not find tomato positivity in osteoblasts and osteocytes even after administration of tamoxifen at P3 and analysis 1 year later (Matsushita et al., 2020).

We, therefore, concluded that the bone growth abnormalities observed in SMN2 1-copy mutants are due to problems in endochondral ossification caused by chondrocyte defects and not due to other Prrx1-lineage skeletal cells.

(b) According to the reviewer's suggestion, we evaluated cell proliferation in the new Figure 1J-L by performing immunostaining for the Ki67 proliferation marker in growth plates.

(d) As the reviewer pointed out, we enhanced the mechanism study and found the reduction of chondrocyte-derived IGF signaling and hypertrophic marker in new Figure 2. We evaluated the density of osteoblasts and osteoclasts, which can affect bone mineralization. We highlighted the limited impact of BMSCs on bone growth in the first two weeks of life. In a previous study, SMN-deleted osteoblasts did not show any issues with ossification (Hensel et al., 2020). In fact, osteoblast density in the SMN2 1-copy mutant was not different from the control, indicating that the skeletal abnormalities can largely be attributed to deficiencies in endochondral ossification caused by chondrocytes. Since chondrocytes are the local source of IGF and our mutants exhibit phenotypes similar to mouse models with reduced IGF, such as downregulated expression of Igf1 and Igfbp3, downregulated IGF-induced hypertrophic gene expression, reduced AKT phosphorylation, proliferation, and growth plate zone length, SMN-deleted chondrocytes probably showed these phenotypes due to decreased IGF secretion. Now, we added new Figure 2A-C, and E.

1. Is the liver the only organ/tissue that supplied IGF to the chondrocytes or are other lateral plate mesoderm-derived cells potential suppliers? It's not possible to pin SMN deletion in chondrocytes as intrinsic ignoring the other bone cell types that it is depleted from in the Prrx1Cre genetic model.

Recently, Oichi et al. reported that the local IGF source in the growth plate is chondrocytes by in situ hybridization and p-AKT staining (Oichi et al., 2023). When we measured IGF in chondrocytes isolated from articular cartilage, the expressions of Igf1 andIgfbp3 were markedly reduced in chondrocytes with SMN deletion compared to controls (New Figure 2E), suggesting that intrinsic SMN expression in chondrocytes plays an important role in the growth plate.

1. Why is SMN protein being isolated from FAPs to assess levels in the null/SMN2 single copy/double copy mutants when the bone defects are supposed to be a chondrocyte-specific phenotype? This protein expression needs to be confirmed in chondrocytes themselves, and or other Prrx1Cre lineaged skeletal cells.

According to the reviewer’s suggestion, we attempted to evaluate the protein levels in chondrocytes of the SMN2 1-copy mutant. However, we were unable to obtain sufficient numbers of chondrocytes, because of poor proliferation of mutant chondrocytes compared to controls in culture conditions. We could obtain ~10^4 viable cells from 1 mouse of SMN2 1-copy mutant. Therefore, our only options for confirming SMN deletion in chondrocytes were DNA and RNA work. As in the Prrx1-lineage FAPs that the amount of SMN protein correlates with the expression levels of full-length SMN mRNA (Figure 2H-J), we expect that the SMN protein in chondrocytes would be fully depleted due to poor full-length SMN mRNA expression (Figure 2H).

1. Figure 2E should have example images of each type of NMJ characterization.

We revised our figure by adding the example images in new Figure 3E.

1. What are the overall NMJ numbers in the normal formation period? Are these constant into the juvenile period when the authors say the deterioration occurs?

We appreciate the reviewer's constructive comments, and it would be interesting to see if we could see a difference in the total number of NMJs. However, there is one NMJ in every myofiber, and each muscle has hundreds to thousands of myofibers. The technical difficulty of confocal imaging an entire muscle, which can be several millimeters across, precludes experiments that count every NMJ and show a difference. It may be possible to do so by combining clearing and confocal line scanning techniques. In our analysis of the NMJ, the formation of the NMJ in the mutant appears to be normal. Additionally, the number of myofibers seems to be the same, and there may be no difference in the total NMJ number.

1. For transplantation experiments the authors sorted YFP or TOMATO+ cells from the Prrx1Cre mice muscles, but refer to them as FAPs. It is known that other cells including tenocyte-like cells, pericytes, and vascular smooth muscle cells are identified by this reporter line. Staining for TOMATO colocalization with PDGFRA would help to clarify this.

In the method ‘Hindlimb fibro-adipogenic progenitors isolation’ section, we sorted 7AAD–Lin–Vcam–Sca1+ population refers to FAPs. For FAPs transplantation, we also used YFP or TOMATO+ FAPs (7AAD–Lin–Vcam–Sca1+). The ‘FAPs transplantation’ method section did not specify the FAPs population in detail. This has been fixed in the new method. Sca1 (Ly6a) is an effective marker for identifying FAPs within Prrx1-lineage cells, as well as Pdgfra (Leinroth et al., 2022).

1. The authors only compare the SMN2 single copy mutant transplantation to contralateral to show rescue, but how does this compare to overall wt morphology?

According to the reviewer’s constructive comment, we compared them with wild-type morphology (new Figure 7A-D).

1. The asterisks of TOMATO+ in Figure 6A are confusing. FAPs do not usually clump together to form such large plaques and are normally much thinner tendrils. What is the reason for this?

As the reviewer states, FAPs have a fibroblast-like morphology with elongated thinner tendrils. The Figure 6A image in the figure shows a Z-sliced cell body portion of FAP, where the nucleus is located, and it appears blunt. We attached imaged tomato+ FAPs, in which their cell body parts are plaque-like.

Author response image 1
Tomato+ FAPs in muscle.

1. Would transplantation of healthy FAPs after NMJ maturation in SMN mutants still rescue the phenotype? Assessment of this is key for therapy intervention timelines moving forward.

It will be very interesting to see if the phenotype improves after NMJ maturation by healthy FAPs transplantation, but this is a technically difficult experiment to do because we found that FAPs do not implant effectively when injected into naive adult muscle. The transplantation into the adult is sufficiently possible if accompanied by an injury, but this eventually leads to new formation of NMJ again. Thus, it seems impossible to do transplantation experiment after NMJ maturation through general methods. If we discover a method to efficiently rescue SMNs from FAPs or identify a factor that affects FAPs' influence on NMJ, then we may be able to conduct this experiment.

Reference

Hensel, N., Brickwedde, H., Tsaknakis, K., Grages, A., Braunschweig, L., Lüders, K. A., Lorenz, H. M., Lippross, S., Walter, L. M., Tavassol, F., Lienenklaus, S., Neunaber, C., Claus, P., & Hell, A. K. (2020). Altered bone development with impaired cartilage formation precedes neuromuscular symptoms in spinal muscular atrophy. Human Molecular Genetics, 29(16), 2662–2673. https://doi.org/10.1093/hmg/ddaa145

Leinroth, A. P., Mirando, A. J., Rouse, D., Kobayahsi, Y., Tata, P. R., Rueckert, H. E., Liao, Y., Long, J. T., Chakkalakal, J. V., & Hilton, M. J. (2022). Identification of distinct non-myogenic skeletal-muscle-resident mesenchymal cell populations. Cell Reports, 39(6), 110785. https://doi.org/10.1016/j.celrep.2022.110785

Matsushita, Y., Nagata, M., Kozloff, K. M., Welch, J. D., Mizuhashi, K., Tokavanich, N., Hallett, S. A., Link, D. C., Nagasawa, T., Ono, W., & Ono, N. (2020). A Wnt-mediated transformation of the bone marrow stromal cell identity orchestrates skeletal regeneration. Nature Communications, 11(1). https://doi.org/10.1038/s41467-019-14029-w

Oichi, T., Kodama, J., Wilson, K., Tian, H., Imamura Kawasawa, Y., Usami, Y., Oshima, Y., Saito, T., Tanaka, S., Iwamoto, M., Otsuru, S., & Enomoto-Iwamoto, M. (2023). Nutrient-regulated dynamics of chondroprogenitors in the postnatal murine growth plate. Bone Research, 11(1). https://doi.org/10.1038/s41413-023-00258-9

Zhou, B. O., Yue, R., Murphy, M. M., Peyer, J. G., & Morrison, S. J. (2014). Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell, 15(2), 154–168. https://doi.org/10.1016/j.stem.2014.06.008

Reviewer #2

Major comments:

1. Regarding bone deficits - CT analysis of bones should be more comprehensive than Figure 1A shows. How about cross-sections? (a) Are bone phenotypes also age-dependent? (b) PCR was done only for SMA and related proteins (such as IGF). IGF protein in the blood and relevant organs should be studied. Why not include biomarkers of osteoblasts or/and osteoclasts and their regulators? (c)

(a) We appreciate the reviewer’s constructive comment. we added longitudinal section views in new Figure 1A and a description of trabecular bone volume and secondary ossification center in the main text.

(b) Age-dependent evaluation is an important point. By adulthood, the difference between the SMN2 1-copy mutant and the control is much larger, and even at birth there is a slight difference, although not as large as at 2 weeks of age. We focused our phenotyping on bone growth at 2 weeks of age, a time when new bone formation by BMSCs is less influential, when bone growth is primarily driven by endochondral ossification of chondrocytes, and before the defect in theNMJ is primarily manifested.

(c) As the reviewer comments, it is important that IGF are evaluated in tissues other than liver. However, the liver is most likely the source of systemic IGF, as shown by the liver-specific deletion of Igf1 and knockout of Igfals, a protein that forms the IGF ternary complex, which is predominantly expressed in the liver. This resulted in a 90% drop in serum IGF levels and a phenotype of shortened femur length and growth plates in the double KO mice (Yakar et al., 2002).

The local IGF source in the growth plate is chondrocytes confirmed by Igf1 in situ hybridization and p-AKT staining (Oichi et al., 2023). From the In situ hybridization data, we can observe that bone marrow and bone do not express Igf1 at all, but only perichondrium and chondrocytes in the resting zone express Igf1 mRNA. Therefore, we can see that the only supplier of IGF among LPM-derived cells is chondrocytes, and in the new figure 2, we measured IGF pathway expression and AKT phosphorylation in chondrocytes. We have confirmed that the expression of Igf1/Igfbp3 is reduced in chondrocytes with SMN deletion.

To assess serum IGF level, we could not set up this experiment condition during our revision period due to the requirement of administrative procedures for purchasing new apparatuses and the limitation of our research funds. However, as previously stated, there is no difference in the expression of Igf1 and Igfals in the liver, which accounts for 90% of serum IGF levels. Therefore, we did not anticipate significant variations in serum IGF levels.

Evaluation of osteoblasts or osteoclasts was done by section staining due to sampling difficulties for PCR. we assessed osteoblasts and osteoclasts state in new Figure 1-figure supplement 2.

1. What is the relationship between deficits of bone deficits and muscle deficits or even NMJ deficits? Are they inter-related? Is skeletal muscle development also defective in Smn∆MPC mice? Can NMJ deficits result from bone deficits? Or vice versa?

Unfortunately, the reviewer's comments are very difficult to clarify in our study using the Prrx1-cre model. In skeletal muscle development, the myofiber number was not significantly different in our mouse models. A study has shown that inactivating noggin, a BMP antagonist expressed in condensed cartilage and immature chondrocytes, results in severe skeletal defects without affecting the early stages of muscle differentiation (Tylzanowski et al., 2006). Therefore, bone may not have a significant impact on the early development of muscle, but later in postnatal development it may have an impact on motor performance issues. The relationship between bone and NMJ hasn't been studied. The impact of bone defects on motor skill may result in muscle weakness and NMJ problems. In our study, we showed that NMJ deficit rescue by transplantation of FAPs and decreased IGF in chondrocytes, a key source of local IGF. This suggests that the functions of FAPs in NMJ and chondrocytes in bone deficit are crucial, rather than each other's influence.

1. Regarding the rescue experiment, the interpretation of the data should be careful. Evidently, healthy FAPs (td-Tomato positive) were transplanted into TA muscles of 10 days-old SMN2 1-copy SmnΔMPC mice, and NMJs were looked at P56. The control was contralateral TA that was injected with the vehicle. As described above, the data had huge SEM and were difficult to interpret or believe. The control perhaps was wrong if FAPs act by releasing "chemicals" because FAPs from one leg may go to other muscles via blood. Second, if FAPs act via contact, the data shown did not support this. Two red FAPs were shown in Figure 6, one of which was superimposed with a nerve track to one of the three NMJs. This NMJ however did not show any difference to the other two, which did not support a contact mechanism. These rescue data were not convincing.

We appreciate the reviewer’s critical comment, but the reviewer appears to have confused the minimum and maximum range bars in the box-and-whisker plot with the SEM error bar in the bar graph. We apologize for the insufficient description of the figure legends section. We revised them. New Figure 7C, which is a bar graph, has a sufficiently short SEM error bar. In contrast, box-and-whisker plots B and D depict the minimum and maximum range, instead of the SEM, and they are significantly different with a p-value of less than 0.001. If FAPs affect the NMJ via a paracrine factor or ECM with a short range of action, they may rescue the NMJ defect in a non-contact-dependent manner, without affecting the contralateral muscle. Also, the FAPs are heterogeneous, so if only a certain subpopulation rescues, the tomato+ FAP in the figure may not be the rescuing cells.

1. For most experiments, the "n" numbers were too small. 3-5 mice were used for bone characterization. For the NMJ, most experiments were done with 3 mice. It was unclear how many NMJs were looked at. Perhaps due to small n numbers, the SEM values were enormous (for example, in Figure 6).

As with the response to the previous comment, this is due to confusion between box-and-whisker plots and bar graphs, and our data was determined to be significant using the appropriate statistical method.

1. Also for experimental design, some experiments included four genotypes of mice (Fig. 1 J,K) whereas some had only three (Fig.1 A, B, C, D and Fig.3) and others had two (many other figures).

In the first experiments to confirm the phenotypes, we tested the 2-copy mutant, but it was not significantly different from the wild type, and in subsequent experiments, we mainly tested the only 1-copy mutant.

1. What was the reason why mixed muscles were used for NMJ characterization (TA versus EDL)? Why not pick a type I-fiber muscle and a type II-fiber muscle?

We appreciate the constructive comment from the reviewer. Firstly, we conducted a phenotype analysis on the TA muscle. For electrophysiological recording, the EDL muscle should be used for intact nerve with muscle preparation, technically. Additionally, for TEM imaging, EDL was a suitable muscle to locate NMJ positions before TEM processing. Both TA and EDL muscles are adjacent and have similar fiber-type compositions. It would be important to observe in different fiber types of muscles, but when we first identified the phenotype, various types of limb muscles showed similar defects, so we focused on specific muscles.

1. The description of mouse strains was confusing. SMN2 transgenic mice (with different copies) were not described in the methods.

We apologize for the insufficient description of the method section. By crossing mice with the SMN2+/+ homologous allele, SMN2 heterologous mice with only one SMN2 allele are SMN2 1-copy mice (SMN2+/0) and SMN2 homologous mice are SMN2 2-copy mice (SMN2+/+). We revised our manuscript method ‘Animals’ section.

ReferenceOichi, T., Kodama, J., Wilson, K., Tian, H., Imamura Kawasawa, Y., Usami, Y., Oshima, Y., Saito, T., Tanaka, S., Iwamoto, M., Otsuru, S., & Enomoto-Iwamoto, M. (2023). Nutrient-regulated dynamics of chondroprogenitors in the postnatal murine growth plate. Bone Research, 11(1). https://doi.org/10.1038/s41413-023-00258-9

Tylzanowski, P., Mebis, L., and Luyten, F. P. (2006). The noggin null mouse phenotype is strain dependent and haploinsufficiency leads to skeletal defects. Dev. Dyn. 235, 1599–1607. doi: 10.1002/dvdy.20782

Yakar, S., Rosen, C. J., Beamer, W. G., Ackert-Bicknell, C. L., Wu, Y., Liu, J. L., Ooi, G. T., Setser, J., Frystyk, J., Boisclair, Y. R., & LeRoith, D. (2002). Circulating levels of IGF-1 directly regulate bone growth and density. Journal of Clinical Investigation, 110(6), 771–781. https://doi.org/10.1172/JCI0215463

Reviewer #3

1. The authors used Prrx1Cre mouse with floxed Smn exon7(Smnf7) mouse carrying multiple (one or two) copies of the human SMN2 gene. Is it expressed both in chondrocytes and mesenchymal progenitors in the limb?

We appreciate the reviewer's comment. We analyzed the deletion of Smn in chondrocytes and FAPs via Cre using genomic PCR and qRT-PCR, as depicted in new Figure 2. The SMN2 allele, which is expressed throughout the body, can rescue Smn knockout mouse lethality (Monani et al., 2000). Indeed, the short limb length and lethality observed in SMN2 0-copy mutants were mitigated by the presence of multiple copies of SMN2. Therefore, both Chondrocytes and FAPs may express SMN2 transcripts from the transgenic SMN2 allele.

1. Page 10 regarding Fig.2E, please show pretzel-like structure. In Figure 2E, plaque, perforated, open, and branched are shown; however, the pretzel is not shown. The same issue is for the Fig. 3D explanation in the text on page 12.

We appreciate the reviewer's constructive feedback. We included illustrative figures of all types of NMJ characterization, and the branched type is identical to the pretzel type. Therefore, we have replaced ‘branched’ with ‘pretzel’ in our text and revised Figure 3E by incorporating the example images.

1. The explanation of the electrophysiology for Fig.4 in the text on pages 12 and 15 (RRP) is not so convincing for the readers. It is advisable to add TEM data for transplantation if it is not technically difficult.

We appreciate the reviewer's critical feedback. Because we did not measure RRP directly, we removed speculation about the possibility of RRP difference. If observing the active zone with TEM and the docking synaptic vesicle would help quantify RRP, it is technically difficult to obtain images of sufficient quality to distinguish the active zones with our current TEM imaging technique.

1. The authors used the word FAP for 7AAD(-)Lin(-)Vcam(-)Sca1(+). It is recommended to show the expression of PDGFR alpha. Furthermore, as the authors stated in the text, mesenchymal progenitors (FAPs) are heterogeneous. Please discuss this point further. Other reports show at least 6 subpopulations using single-cell analyses (Cell Rep. 2022).

In the report, Ly6a (Sca1) is a good marker for FAPs, as well as Pdgfra (Leinroth et al., 2022). The 6 subpopulations expressed Ly6a. The one of subpopulations associated with NMJ was discovered. This population expressed Hsd11b1, Gfra1, and Ret and is located adjacent to the NMJ and responds to denervation, indicating an increased possibility of interaction with the NMJ organization. In further our study, we aim to determine which subpopulations are crucial for NMJ maturation by transplanting them to mutants for rescue.

1. How do authors determine the number of FAP cells for transplantation?

The FAPs transplantation was performed according to a previously reported our study (Kim et al., 2021).

ReferenceKim, J. H., Kang, J. S., Yoo, K., Jeong, J., Park, I., Park, J. H., Rhee, J., Jeon, S., Jo, Y. W., Hann, S. H., Seo, M., Moon, S., Um, S. J., Seong, R. H., & Kong, Y. Y. (2022). Bap1/SMN axis in Dpp4+ skeletal muscle mesenchymal cells regulates the neuromuscular system. JCI Insight, 7(10). https://doi.org/10.1172/jci.insight.158380

Leinroth, A. P., Mirando, A. J., Rouse, D., Kobayahsi, Y., Tata, P. R., Rueckert, H. E., Liao, Y., Long, J. T., Chakkalakal, J. V., & Hilton, M. J. (2022). Identification of distinct non-myogenic skeletal-muscle-resident mesenchymal cell populations. Cell Reports, 39(6), 110785. https://doi.org/10.1016/j.celrep.2022.110785

Monani, U. R., Sendtner, M., Coovert, D. D., Parsons, D. W., Andreassi, C., Le, T. T., Jablonka, S., Schrank, B., Rossol, W., Prior, T. W., Morris, G. E., & Burghes, A. H. M. (2000). The human centromeric survival motor neuron gene (SMN2) rescues embryonic lethality in Smn(-/-) mice and results in a mouse with spinal muscular atrophy. Human Molecular Genetics, 9(3), 333–339. https://doi.org/10.1093/hmg/9.3.333

https://doi.org/10.7554/eLife.92731.3.sa4

Article and author information

Author details

  1. Sang-Hyeon Hann

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing
    Contributed equally with
    Seon-Yong Kim
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-0871-3414
  2. Seon-Yong Kim

    Department of Physiology, Dental Research Institute, Seoul National University School of Dentistry, Seoul, Republic of Korea
    Contribution
    Data curation, Formal analysis, Validation, Investigation, Methodology, Writing – review and editing
    Contributed equally with
    Sang-Hyeon Hann
    Competing interests
    No competing interests declared
  3. Ye Lynne Kim

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Validation, Investigation, Writing – review and editing
    Competing interests
    No competing interests declared
  4. Young-Woo Jo

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Data curation, Formal analysis, Methodology, Writing – review and editing
    Competing interests
    No competing interests declared
  5. Jong-Seol Kang

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Formal analysis, Investigation, Methodology
    Competing interests
    No competing interests declared
  6. Hyerim Park

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Investigation, Visualization, Methodology
    Competing interests
    No competing interests declared
  7. Se-Young Choi

    Department of Physiology, Dental Research Institute, Seoul National University School of Dentistry, Seoul, Republic of Korea
    Contribution
    Supervision, Funding acquisition, Methodology, Project administration, Writing – review and editing
    For correspondence
    sychoi@snu.ac.kr
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7534-5167
  8. Young-Yun Kong

    School of Biological Sciences, Seoul National University, Seoul, Republic of Korea
    Contribution
    Supervision, Funding acquisition, Methodology, Project administration, Writing – review and editing
    For correspondence
    ykong@snu.ac.kr
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-7335-3729

Funding

National Research Foundation of Korea (NRF-2022R1A2C3007621)

  • Young-Yun Kong

National Research Foundation of Korea (NRF-2020R1A5A1018081)

  • Young-Yun Kong

National Research Foundation of Korea (NRF-2020R1A2C3011464)

  • Se-Young Choi

The funders had no role in study design, data collection, and interpretation, or the decision to submit the work for publication.

Acknowledgements

We express our gratitude to the Kong and Choi laboratory members for their valuable feedback during the project. This work was supported by the National Research Foundation of Korea (NRF-2022R1A2C3007621, NRF-2020R1A5A1018081, and NRF-2020R1A2C3011464).

Ethics

The care and treatment of animals in this study were approved by the Institutional Animal Care and Use Committee (IACUC) protocols (SNU-210313-1) of Seoul National University.

Senior Editor

  1. Timothy E Behrens, University of Oxford, United Kingdom

Reviewing Editor

  1. Vitaly Ryu, Icahn School of Medicine at Mount Sinai, United States

Version history

  1. Sent for peer review: September 21, 2023
  2. Preprint posted: October 10, 2023 (view preprint)
  3. Preprint posted: November 7, 2023 (view preprint)
  4. Preprint posted: January 23, 2024 (view preprint)
  5. Version of Record published: February 6, 2024 (version 1)

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You can cite all versions using the DOI https://doi.org/10.7554/eLife.92731. This DOI represents all versions, and will always resolve to the latest one.

Copyright

© 2023, Hann, Kim et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Sang-Hyeon Hann
  2. Seon-Yong Kim
  3. Ye Lynne Kim
  4. Young-Woo Jo
  5. Jong-Seol Kang
  6. Hyerim Park
  7. Se-Young Choi
  8. Young-Yun Kong
(2024)
Depletion of SMN protein in mesenchymal progenitors impairs the development of bone and neuromuscular junction in spinal muscular atrophy
eLife 12:RP92731.
https://doi.org/10.7554/eLife.92731.3

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